We thank Pierre G&#246;nczy and the BioImaging &amp; Optics Platform (BIOP) at &#201;cole Polytechnique F&#233;d&#233;rale de Lausanne (EPFL), Lausanne, Switzerland, where the SIM images of the centrioles were acquired. Nikolai Klena and Davide Gambarotto are supported by the European Research Council (ERC) Starting Grant (StG) 715289 (ACCENT) and Maeva Le Guennec, Paul Guichard, and Virginie Hamel by the Swiss National Science Foundation (SNSF) PP00P3_157517. Susanne Borgers is supported by the University of Geneva.

<ol>
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The authors have nothing to disclose.

One of the challenges in biology is to decipher the precise localization of proteins in an architectural context. The centriole is an ideal structure to apply these methods, as its architecture has been studied using cryo-electron tomography, revealing interesting ultrastructural features along its length. However, due to its dimensions close to the resolution limit in optical microscopy, it is difficult to precisely localize a protein by immunofluorescence to a structural sub-region of the centriole using conventional microscopes30.
The resolution in optical microscopy is limited by the diffraction of light that gives, roughly, a lateral maximum resolution of 200 nm in optical microscopy24. However, this limit has been by-passed by one of the major breakthroughs of the last 20 years in optical microscopy: the invention of super-resolution methods. These approaches can image beyond the diffraction limits at different resolutions: 120 nm for SIM, about 50 nm for stimulated emission depletion (STED), and 20&#8211;40 nm for single-molecule localization microscopy (SMLM)24. With these new developments of super-resolution microscopy, the structural sub-regions of the centriole are attainable. However, in practice, it is still difficult to accurately determine the localization of a protein to a structural element for the main reason that the mature centrioles exist in only 2 copies per cell and have random orientations, which makes the interpretation of localization difficult. For this reason, a protocol was set up that allows researchers to image by super-resolution a large number of centrioles, increasing the chance to observe non-ambiguous orientations. Importantly, as this method relies on the use of isolated centrioles, we are providing a method to purify intact C. reinhardtii centrioles that contain mature centrioles and procentrioles.
Finally, owing to the range of centriole orientations that can be imaged with this protocol, single-particle analysis can be applied using electron microscopy software. This results in the generation of average classes of centrioles in a particular orientation. Importantly, these resulting 2-D images can then be used to assess the localization of a specific protein along the centriole. Indeed, this method can be applied to dual-color super-resolution images, and one color can be used to reveal the centriole skeleton (e.g., tubulin), while the other color can be attributed to a specific centriolar protein. By subtracting the averages obtained with one color or two colors, it becomes easier to register a protein along the centriole (proximal, central, or distal). Note that the two channels should be accurately aligned to prevent any misleading interpretations. Moreover, averages of top views will help decipher if a protein localizes inside the centriolar lumen, along the microtubule wall, or outside the centriole.
This method has the advantage to ascertain the localization of specific proteins that could be difficult to localize otherwise due to heterogeneous labeling. Note that other methods to map proteins within the centrioles have been described in correlative 3-D SIM/SMLM with, for instance, assessing the specific orientations of the centrioles by determining the elliptical profile of a marker forming a torus around the centriole by SIM imaging. Using this parameter, it is possible to localize protein with a precision of 4&#8211;5 nm30. Note also that the method described here uses isolated centrioles with intact procentrioles, a sign that the centriole architecture is most likely largely conserved. However, we cannot exclude that some architectural features are disturbed during purification, such as the centriole diameter varying with the concentration of divalent cations as amplified with the isolation of the human centrosome5.
One of the critical steps of the protocol presented here is to obtain sufficiently concentrated isolated centrioles in different orientations amenable to Fluo-SPA. To do so, first ensure the purity and the efficiency of the centriole isolation procedure. A low concentration of isolated centrioles will prevent proper imaging and further image processing. For this purpose, we are providing a method to enrich the number of centrioles per field of view. Depending on the number of centrioles in the fraction used, the volume loaded in the concentrator should be adjusted, with a maximum volume of 250 &#181;L.
Importantly, this method has been developed for a cell wall minus cw15- C. reinhardtii cells. In this strain, the fragility of the cell wall allows for a proper lysis of the cells and, thus, the liberation of its content. This protocol is not efficient for wild-type C. reinhardtii cells, as the cell wall prevents a proper lysis. Alternative strategies such as sonication or a pre-incubation of the cells with autolysin, an enzyme that can degrade the cell wall31, would have to be put in place to alter the cell wall prior to applying the isolation protocol presented here.
This set-up can be used with different types of microscopes, ranging from conventional confocal microscopes to high throughput dedicated super-resolution microscopes. Note that when doing SMLM, a special buffer is required for proper imaging and, thus, a chamber adapted for a 12 mm coverslip with the buffer on top of the coverslip should be used. Subsequent imaging will be performed with inverted microscopes. If the microscope set-up does not allow a 12 mm coverslip, the protocol presented here can be applied to 18 mm coverslips using a 30 mL round-bottom tube and a modified adaptor and concentrator. It is also important to note that the quality of the final reconstruction in SMLM will depend on the quality of the staining and of the primary antibody used, as well as the method of fixation.
In summary, we are providing a method that can be applied to image numerous centrioles followed by Fluo-SPA that will generate averages of centrioles in different orientations, thus helping to localize centriolar protein with precision. Importantly, this method can be applied more generally to isolated centrioles from other species, to other organelles, or to large macromolecular assemblies. Finally, the sample preparation approach presented here, combined with recent algorithm development for single-particle analysis of fluorescent SMLM data32, could open further improvement in the molecular cartography of large macromolecular assemblies.

The centriole is an evolutionarily conserved organelle that lies at the core of the centrosome in animal cells and can act as a basal body (referred to as centrioles hereafter) to template cilia or flagella in many eukaryotes1,2. As such, centrioles are critical for fundamental cell biological processes ranging from spindle assembly to cell signaling. Therefore, defects in centriole assembly or function have been associated with several human pathologies including ciliopathies and cancers3.
Centrioles are nine-fold, symmetrical, microtubule triplet-based cylindrical structures that are, typically, ~450 nm long and ~250 nm wide4,5,6,7. Conventional electron microscopy and cryo-electron tomography of centrioles from different species have revealed that centrioles are polarized along their long axis with three distinct regions: a proximal region, a central core, and a distal region5,7,8,9,10,11. Importantly, each of these regions displays specific structural features. First, the lumen of the 100 nm-long proximal region contains the cartwheel structure connected to the microtubule triplet through the pinhead element12. Second, the 300&#8211;400 nm-long central region contains fibrous densities in the lumen and structural features along the inner face of the microtubules: the Y-shaped linker, the C-tubule tail, and the A-tubule stub9. Finally, the 50&#8211;100 nm distal region exhibits sub-distal and distal appendages that surround the distal part of the centriole5,13.
The last two decades have been marked by the discovery of an increasing number of centriolar proteins, leading to a current estimation of about 100 distinct proteins being part of the centriole14,15,16,17. Despite these advances, the precise localization of these proteins within the centriole remains elusive, particularly within structural sub-regions. Importantly, assigning a precise localization to structural regions of the centriole is crucial for a better understanding of their function. In this respect, C. reinhardtii centrioles have been instrumental in both aspects by first delimitating the different structural features along the cylinder9,18,19, which then allowed researchers to correlate the localization of a subset of proteins using fluorescent microscopy to a sub-structural region. This includes, for example, the proteins Bld12p and Bld10p, which localize in the proximal region, and in the cartwheel structure in particular20,21,22,23. The list of substructure localized proteins also includes POB15 and POC16, two novel proteins identified by mass spectrometry that decorate the inner central core region of C. reinhardtii centrioles17.
This paper provides a complete description of the method developed to isolate and image C. reinhardtii centrioles for subsequent super-resolution microscopy and single-particle averaging. To achieve this goal, it is important to delineate the technical limitations that need to be overcome. First, centriole purification can affect the overall architecture, with the cartwheel structure often being lost during the various steps of isolation9. Secondly, the dimensions of the centriole are very close to the diffraction limit in optical microscopy. Indeed, the lateral resolution that can be obtained in confocal microscopy is around 200 nm24, similar to the diameter of the centriole, and the resolution in the z-axis is about 2&#8211;3x lower, leading to an anisotropic volume. Thirdly, the heterogeneity of antibody labeling and centriole orientation could limit the interpretation needed to localize a protein in a specific centriolar sub-region. Finally, centrioles exist in only two copies per cell, making it difficult to acquire a large number of images and find an unambiguous centriole orientation. To circumvent these technical issues, we developed a method that relies on applying super-resolution microscopy on large numbers of isolated centrioles that adopt various orientations. We will first describe a protocol to purify C. reinhardtii centrioles that enables the purification of structurally intact centrioles and procentrioles containing the cartwheel. Then, we will describe a step-by-step protocol to concentrate the centrioles on coverslips for imaging by conventional or super-resolution fluorescent microscopy. This important step allows for increasing the number of centrioles imaged in multiple orientations. Finally, we will describe a procedure to perform single-particle averaging on data acquired on fluorescent microscopes that facilitates the detection of centrioles in different orientations. Altogether, this method can be applied to image centrioles from various species or other large macromolecular assemblies.

<table><tbody><tr><td>Mouse monoclonal anti-apha tubulin (clone DM1A)</td><td>Abcam</td><td>ab7291&amp;nbsp;</td><td>dilution 1:300</td></tr><tr><td>DNaseI</td><td>Roche</td><td>10104159001</td><td></td></tr><tr><td>12 mm coverslips</td><td>Roth</td><td>YX03.1</td><td></td></tr><tr><td>18 mm coverslips</td><td>Roth</td><td>LH23.1</td><td></td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fluka</td><td>60355</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fluka</td><td>60230</td><td></td></tr><tr><td>Tris base</td><td>Biosolve Chimie SARL</td><td>0020092391BS</td><td></td></tr><tr><td>acetic acid</td><td>Carlo Erba Reagents</td><td>524520</td><td></td></tr><tr><td>NH&lt;sub&gt;4&lt;/sub&gt;Cl</td><td>Sigma</td><td>A-4514</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C-7902</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>63140-500G-F</td><td></td></tr><tr><td>steritop filter</td><td>Millipore</td><td>SCGPT05RE</td><td></td></tr><tr><td>sucrose</td><td>Sigma</td><td>S7903-1KG</td><td></td></tr><tr><td>HEPES</td><td>AppliChem PanReac</td><td>A3724,0250</td><td></td></tr><tr><td>PIPES</td><td>Sigma</td><td>P6757-500G</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>ACROS ORGANICS</td><td>197530010</td><td></td></tr><tr><td>NP-40</td><td>AppliChem PanReac</td><td>A1694,0250</td><td></td></tr><tr><td>Round-bottom (Kimble) tubes 15 mL</td><td>Fisherscientific</td><td>09-500-34</td><td></td></tr><tr><td>Round-bottom (Kimble) tubes 30 mL</td><td>Fisherscientific</td><td>09-500-37</td><td></td></tr><tr><td>cover glass staining rack</td><td>Thomas scientific</td><td>8542E40</td><td></td></tr><tr><td>crystal polystyrene transmission lab box</td><td>FISHERS</td><td>11712944</td><td></td></tr><tr><td>Methanol</td><td>VWR</td><td>20864.32</td><td></td></tr><tr><td>BSA</td><td>Roche</td><td>10735086001</td><td></td></tr><tr><td>Triton X100</td><td>Roth</td><td>3051.3</td><td></td></tr><tr><td>goat anti-mouse coupled to Alexa 488&amp;nbsp;</td><td>invitrogen</td><td>A11029</td><td>dilution 1:1,000</td></tr><tr><td>goat anti-rabbit coupled to Alexa 568</td><td>invitrogen</td><td>A11036</td><td>dilution 1:1,000</td></tr><tr><td>mounting medium</td><td>abcam</td><td>ab188804</td><td></td></tr><tr><td>Tube, thinwall polypropylene</td><td>Beckman Coulter</td><td>326823</td><td></td></tr><tr><td>Poly-D-Lysine 1 mg/mL</td><td>SIGMA&amp;nbsp;</td><td>A-003-E</td><td></td></tr><tr><td>Mouse monoclonal anti-Polyglutamylation modification mAb (GT335)</td><td>Adipogen</td><td>AG-20B-0020</td><td>dilution 1:1,000</td></tr><tr><td>glycerol mounting medium with DAPI and DABCO</td><td>Abcam</td><td>ab188804</td><td></td></tr><tr><td>50 mL conical tubes</td><td>Falcon</td><td>14-432-22</td><td></td></tr><tr><td>Eppendorf 5810R centrifuge</td><td>Eppendorf</td><td>5811000622</td><td></td></tr><tr><td>Beckman JS-13.1 swinging bucket rotor</td><td>Beckman Coulter</td><td>346963</td><td></td></tr><tr><td>Beckman SW 32Ti rotor</td><td>Beckman Coulter</td><td>369694</td><td></td></tr><tr><td>parafilm</td><td>Bemis&amp;nbsp;</td><td>13-374-10</td><td></td></tr><tr><td>Leica TCS SP8 with Hyvolution mode</td><td>Leica&amp;nbsp;</td><td></td><td></td></tr><tr><td>OSRAM L18W/954 LUMILUX&amp;nbsp;</td><td>Luxe Daylight/ OSRAM</td><td></td><td></td></tr><tr><td>Whatman filter paper</td><td>Sigma&amp;nbsp;</td><td>WHA1001325</td><td></td></tr><tr><td>CrSAS-6/Bld12 antibody</td><td></td><td></td><td>dilution 1:300 (Hamel &lt;em&gt;el al.&lt;/em&gt;, 2014)</td></tr><tr><td>Scipion</td><td>http://scipion.i2pc.es/</td><td></td><td></td></tr><tr><td>EM CCD camera (Andor iXON DU897)</td><td>Andor</td><td></td><td></td></tr></tbody></table>

isolation and fluorescence imaging for single-particle reconstruction of chlamydomonas centrioles

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell motility, or cell signaling. The green algae Chlamydomonas reinhardtii has proven to be an insightful model in the study of centriole architecture, function, and protein composition. Despite great advances toward understanding centriolar architecture, one of the current challenges is to determine the precise localization of centriolar components within structural regions of the centriole in order to better understand their role in centriole biogenesis. A major limitation lies in the resolution of fluorescence microscopy, which complicates the interpretation of protein localization in this organelle with dimensions close to the diffraction limit. To tackle this question, we are providing a method to purify and image a large number of C. reinhardtii centrioles with different orientations using super-resolution microscopy. This technique allows further processing of data through fluorescent single-particle averaging (Fluo-SPA) owing to the large number of centrioles acquired. Fluo-SPA generates averages of stained C. reinhardtii centrioles in different orientations, thus facilitating the localization of distinct proteins in centriolar sub-regions. Importantly, this method can be applied to image centrioles from other species or other large macromolecular assemblies.

C. reinhardtii Centriole Isolation: 
To isolate centrioles, cw15- C. reinhardtii cells were grown in liquid culture for several days under light and subsequently pelleted by centrifugation at 600 x g for 10 min. Pelleted cells were washed 1x with PBS and resuspended in a deflagellation buffer prior to deflagellation by performing a pH shock using 0.5 M acetic acid to a final pH of 4.5&#8211;4.7 for 2 min (Figure 1A). An addition of 1 N KOH was used to restore pH to 7.0. To separate detached flagella from cell bodies, deflagellated cells were first centrifuged to remove the bulk of the flagella. The pellet was then washed 2x with PBS and resuspended in 30&#160;mL of PBS prior to loading it slowly on a 25%-sucrose cushion (Figure 1B). The tube was spun at 600 x g for 15 min at 4 &#176;C to remove most of the detached flagella. After centrifugation, the cell bodies were spread in the sucrose cushion (Figure 1C) and recovered by removing the approximately 30 mL of supernatant (until the red arrow in Figure 1C). The resulting 20 mL of washed cells were then resuspended in 20 mL of cold PBS and centrifuged at 600 x g for 10 min. Next, the supernatant was discarded, and the cells were completely resuspended in 10 mL of PBS. The cells were transferred to a 250&#160;mL bottle and the lysis buffer was added to the resuspended cells at once. DNase was added to the lysis and incubated for 1 h at 4 &#176;C. After a centrifugation step to remove cell debris (see the white pellet in Figure 1D), the supernatant was collected and carefully loaded onto a 2 mL 60%-sucrose cushion prior to centrifugation at 10,000 x g for 30 min at 4 &#176;C. Note that for 100 mL of lysis, 8 tubes of 15 mL were used to perform this step, corresponding to 12.5 mL of lysis buffer loaded on 2 mL of sucrose per tube. After centrifugation, most of the supernatant was removed up to 1 mL above the cushion. The 1 mL of remaining supernatant was then collected with the 2 mL cushion and then mixed and pooled to obtain a final volume of 24 mL. The pool was then loaded on a 40%-, 50%-, 70%-sucrose gradient and spun at 68,320 x g for 1 h and 15 min at 4 &#176;C. Finally, the isolated centrioles were collected by making a hole in the bottom part of the centrifugation tube using a needle and the drops were collected in 12 x 500 &#181;L fractions. Due to the high density of the different sucrose, the drop formed very slowly at the beginning (70% sucrose) and then more rapidly (40% sucrose).
Immunofluorescence of Isolated Centrioles 
To assess the quality of the isolation procedure, 10 &#181;L of each collected gradient fraction was then centrifuged onto a coverslip using a coverslip support adaptor (Figure&#160;2A-2D). Importantly, to remove safely the coverslip, a custom hooked device was designed (Figure 2B). Next, the coverslips were analyzed by immunofluorescence. Antibodies against CrSAS-6(Bld12p) were used in this study to indicate the presence of the cartwheel structure and &#945;-tubulin (DMA1) to highlight the centriolar wall. By counting the number of centrioles that were positive for both CrSAS-6 and &#945;-tubulin per field of view and then calculating the total number of centrioles per fractions, it was possible to determine which fractions were enriched for isolated centrioles (Figure&#160;2F-2H). Interestingly, 6 fractions were enriched for centrioles (Figure&#160;2F and 2G, fractions #1&#8211;6), with a peak for fraction #3, while the last fractions were not, indicating that the purification worked. Note that in this particular experiment, 95% of the total centrioles were positive for CrSAS-6 and &#945;-tubulin in fraction #3. This indicates that most isolated centrioles retained their cartwheels. If no enrichment of centrioles is observed in the first fractions, the isolation procedure did not work and should be repeated. Note that some flagellar pieces can be observed mostly in the fractions devoid of centrioles.
Next, to calculate the total number of centrioles per &#181;L, the number of centriole per field of view should be multiplied by the ratio as presented in Figure 2H. The resulting number should then be divided by the volume of the fraction used for the immunofluorescence to obtain the number of centrioles per &#181;L. In this particular isolation procedure, the most enriched fraction contained about 37 centrioles for an area of 0.00846 mm2 (with a field of view of 92 x 92 &#181;m2). The surface of the tube section was 7.5 mm in radius with a total surface of 176 mm2. The corresponding ratio was then 176/0.00846 = 20,803.8, so a total of 769,740 centrioles (37x20,803.8) in 10 &#181;L. Therefore, the number of centrioles in 1 &#181;L was 76,974.
Concentration of Isolated Centrioles on Coverslips: 
Increasing the number of centrioles per field increases the chance of detecting unambiguous centriole orientation, as well as increasing the chance of detecting similar orientations that can be used for further particle averaging procedures. As centrioles from the concentrated fractions are still sparse on the coverslip, we developed a centrifugation accessory to concentrate centrioles in the middle of the coverslip (Figure 3A) named a concentrator. Note that a .stl file with the precise measures for 3-D printing is provided with this manuscript.
First, one coverslip of 12 mm was mounted on the concentrator (Figure 3B). The adaptor was placed on top of the coverslip and the round-bottom tube was inverted and placed over the assembled adaptor and concentrator. The round-bottom tube was then gently inverted, thus allowing the loading of the sample (Protocol step 4.2). The centrioles were then centrifuged at 10,000 x g for 10 min at 4 &#176;C. After that, the centrioles were subjected to immunofluorescence and stained for CrSAS-6/Bld12p and &#945;-tubulin (Figure&#160;3C-3E). Importantly, without concentrator, about 30 centrioles were seen per field of view (Figure 3C), whereas 183 centrioles were seen per field of view when the concentrator was used (Figure&#160;3D-3F). Note that the centrioles covered only a disk of 4 mm in diameter in the middle of the coverslip. This result demonstrates that the concentration step works and allows a 6-fold enrichment of centrioles in a defined region of the coverslips, thus easing their detection and imaging.
Fluorescent Single-particle Averaging of Isolated C. reinhardtii Centrioles: 
Here, using SIM microscopy that can reach a resolution of about 120 nm, centrioles stained for monoglutamylated tubulin (GT335, Figure 4), a tubulin modification present in centriolar microtubules, were imaged24. C. reinhardtii centrioles are about 500 nm long, always in pairs, and often found with associated, newly duplicated probasal bodies (referred to as procentrioles hereafter) and striated microtubule-associated fibers19. Therefore, this final assembly was about 1 &#181;m large. For this reason, and in order to image the centrioles in their entirety, we recommend acquiring a Z-stack on the isolated centrioles.
Here, after the acquisition, a final image was generated by performing a maximum intensity projection using ImageJ28 (image/Stacks/Z project/max intensity projection, Figure 4B). From such images, a single-particle analysis using cryo-electron microscopy software was performed to make classes of centrioles with similar orientations, and then averaging was performed. To do so, the image color was first inverted so as to better visualize the objects (Figure 4C). Centrioles were picked manually in a box centered over each particle using the freely available Scipion software29 that integrates several electron microscopy software programs such as Xmipp3 (Figure 4D). Note that the size of the box has to be defined by the user. Here, boxes of 50 x 50 pixels for a pixel size of 31.84 nm were used. Next, all particles were extracted (Figure 4E) and aligned using Xmipp3 (Figure 4F). Next, a circular mask of 12 pixels in radius was applied to isolate each centriole from the centriolar-pair (Figure 4G). The particles were then classified, using Xmipp3, to generate several averages. Only homogenous class averages were kept, meaning that particles that diverge from the class averages were manually excluded. This step was repeated in order to generate a near-perfect average for each chosen orientation. After five iterations, two classes of averages were generated: a top view from 63 objects (Figure 4H) and a side view from 98 particles (Figure 4I) of monoglutamylated centrioles. The dimensions of the object were determined by measuring the distance between peaks of the intensity plot profile along the monoglutamylation signal.
Importantly, the length of the side-view class average is 260 nm with a diameter of 250 nm, comparable to the measured monoglutamylated tubulin signal that was shown to localize to a region of 286 &#177; 33 nm in length within the core of the centriole17.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58109/58109fig1.jpg" /> 
Figure 1:&#160;Purification of C. reinhardtii&#160;centrioles. (A) This is a schematic representation of each step leading to the isolation of C. reinhardtii centrioles. It includes representative steps of the protocol (B) before and (C) after centrifugation onto the 25%-sucrose gradient. The red arrow in panel C indicates the minimum volume to keep after centrifugation. (D) This panel shows the white pellet of lysed cells after centrifugation. The black arrow indicates the pellet. <a href="https://www.jove.com/files/ftp_upload/58109/58109fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58109/58109fig2.jpg" /> 
Figure 2:&#160;Centrifugation set-up to perform immunofluorescence on isolated centrioles. (A) 10 &#181;L of each collected fraction is diluted first in 100 &#181;L of 10mM K-PIPES (pH 7.2). (B) This is a schematic representation of the centrifugation devices, encompassing a 15 mL round-bottom tube, a 12 mm coverslip, an adaptor for the coverslip, and a custom-made hooked device to recover the coverslip after centrifugation. Panels C and D show drawings explaining how to recover the coverslip after centrifugation. (C) Place the hooked tool into the hole present in the slotted edge of the adaptor and (D) gently pull. (E) These are pictures of the humid chamber needed to perform immunofluorescence imaging. The arrow indicates the 12 mm coverslip. (F) These are representative confocal images at 63X (zoom 2) of gradient fractions #1-8, collected during the purification procedure and stained for CrSAS-6 (red) and &#945;-tubulin (green). The insets correspond to the region indicated by a white box in the lower magnification views. Scale bar = 10 &#181;m. (G) This is a graph representing the number of centrioles positive for both CrSAS-6 and &#945;-tubulin per field of view in each fraction. Note the enrichment in fractions #1-6. The average number of centrioles per field in each fraction: #1 = 16.3 &#177; 4.7, #2 = 24.0 &#177; 4.6, #3 = 37.0 &#177; 17.0, #4 = 23.3 &#177; 11.4, #5 = 14.3 &#177; 2.1, #6 = 13.7 &#177; 9.3, #7 = 2.7 &#177; 1.2, #8 = 1.0 &#177; 0.0, #9 = 1.0 &#177; 0.0, #10 = 0.0 &#177; 0.0, #11 = 0.3 &#177; 0.6, #12 = 0.0 &#177; 0.0. For each fraction, 3 random fields were imaged. Note that, when counting the most enriched fraction #3, we found that 95% of the centrioles are positive for CrSAS-6 and &#945;-tubulin (n = 205 centrioles). (H) A ratio of the number of centrioles present in the measured area of the micrographs is used to calculate the total number of centrioles present in the area of the tube. The centriole number per &#181;L can be calculated from the 10 &#181;L originally centrifuged fractions. <a href="https://www.jove.com/files/ftp_upload/58109/58109fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58109/58109fig3.jpg" /> 
Figure 3: Isolated centrioles are centrifuged using concentrators. (A) This panel shows the centrifugation devices needed to concentrate centrioles onto coverslips prior to immunofluorescence, including a 15 mL round-bottom tube, a concentrator, a 12 mm coverslip, and a support adaptor apparatus. (B) This panel shows the steps to assemble the centrifugation device. Panels C-E show confocal images of centrioles stained for CrSAS-6 (red) and &#945;-tubulin (green) (C) without or (D-E) with the concentrator. The insets correspond to the region indicated by a white box in the lower magnification views. The scale bar is 10 &#181;m. Note that centrioles are enriched in the middle of the coverslip (the dotted line represents the border of the concentrated area). (F) This graph represents the number of centrioles per field of view without and with the concentrator. Five random fields of view were analyzed. The average number of centrioles is, without concentrator, 29.8 &#177; 2.9, and with concentrator, 182.6 &#177; 11.5, P &#60;0.0001. The statistical significance was assessed by an unpaired t-test. <a href="https://www.jove.com/files/ftp_upload/58109/58109fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58109/58109fig4.jpg" /> 
Figure 4:&#160;Single-particle averaging on isolated C. reinhardtii centrioles. (A) This panel shows Z stack images of centrioles stained with GT335 and acquired using a SIM microscope. (B) These panels show a maximal intensity Z projection of the stacked images. Scale bar = 1 &#181;m. (C) These panels show a representative image with an inverted contrast. The inset represents a zoom-in to visualize the centrioles better. (D) These panels show particle picking. The inset indicates how the particles were picked. (E) These are examples of 5 extracted particles. (F) This panel shows the particles after alignment. (G) This panel shows the particles after applying a mask. Panels H and I show two class averages: (H) a top view (63 particles) and (I) a side view (98 particles). The double arrows indicate the dimensions of the GT335 signal. Scale bar = 250 nm. <a href="https://www.jove.com/files/ftp_upload/58109/58109fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1.&#160;<a href="https://www.jove.com/files/ftp_upload/58109/Adaptor_coverslip.stl" target="_blank">Please click here to download this file.</a>
Supplementary File 2.&#160;<a href="https://www.jove.com/files/ftp_upload/58109/concentrator.stl" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles at JoVE.com

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

We have developed a strategy to purify and image a large number of centrioles in different orientations amenable for super-resolution microscopy and single-particle averaging.

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell ...

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9.4K Views.  University of Geneva. We have developed a strategy to purify and image a large number of centrioles in different orientations amenable for super-resolution microscopy and single-particle averaging.

Video: Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

Immunology and Infection

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, however, the lateral resolution of super-resolution imaging is only ~100 nm. The low resolution is mainly due to the high autofluorescence background of photosynthetic cells caused by high-intensity lasers that are required for super-resolution imaging, such as stochastic optical reconstruction microscopy (STORM). Here, we describe a photobleaching-assisted STORM method which was developed recently for imaging the marine picocyanobacterium Prochlorococcus. After photobleaching, the autofluorescence of Prochlorococcus is effectively reduced so that STORM can be performed with a lateral resolution of ~10 nm. Using this method, we acquire the in vivo three-dimensional (3-D) organization of the FtsZ protein and characterize four different FtsZ ring morphologies during the cell cycle of Prochlorococcus. The method we describe here might be adopted for the super-resolution imaging of other photosynthetic organisms.

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Here we describe a photobleaching method to reduce the autofluorescence of cyanobacteria. After photobleaching, stochastic optical reconstruction microscopy is used to obtain three-dimensional super-resolution images of the cyanobacterial FtsZ ring.

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, ...

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to facilitate sensory functions in cells. The level of a protein may be regulated differently at centrosomes than at other .cellular locations, and the variation in the centrosomal level of several proteins at different points of the cell cycle appears to be crucial for the proper regulation of centriole assembly. We developed a quantitative fluorescence microscopy assay that measures relative changes in the level of a protein at centrosomes in fixed cells from different samples, such as at different phases of the cell cycle or after treatment with various reagents. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region, and normalize that measurement against the same for another protein that does not vary under the chosen experimental condition. Utilizing this assay in combination with BrdU pulse and chase strategy to study unperturbed cell cycles, we have quantitatively validated our recent observation that the centrosomal pool of VDAC3 is regulated at centrosomes during the cell cycle, likely by proteasome-mediated degradation specifically at centrosomes.

Here, a novel quantitative fluorescence assay is developed to measure changes in the level of a protein specifically at centrosomes by normalizing that protein&#8217;s fluorescence intensity to that of an appropriate internal standard.

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to ...

Immunofluorescence Analysis of Endogenous and Exogenous Centromere-kinetochore Proteins

"Centromeres" and "kinetochores" refer to the site where chromosomes associate with the spindle during cell division. Direct visualization of centromere-kinetochore proteins during the cell cycle remains a fundamental tool in investigating the mechanism(s) of these proteins. Advanced imaging methods in fluorescence microscopy provide remarkable resolution of centromere-kinetochore components and allow direct observation of specific molecular components of the centromeres and kinetochores. In addition, methods of indirect immunofluorescent (IIF) staining using specific antibodies are crucial to these observations. However, despite numerous reports about IIF protocols, few discussed in detail problems of specific centromere-kinetochore proteins.1-4 Here we report optimized protocols to stain endogenous centromere-kinetochore proteins in human cells by using paraformaldehyde fixation and IIF staining. Furthermore, we report protocols to detect Flag-tagged exogenous CENP-A proteins in human cells subjected to acetone or methanol fixation. These methods are useful in detecting and quantifying endogenous centromere-kinetochore proteins and Flag-tagged CENP-A proteins, including those in human cells.

Here we report protocols to detect endogenous and exogenous centromere-kinetochore proteins in human cells and quantify these protein levels at centromeres-kinetochores by indirect immunofluorescent staining through the use of fixation (paraformaldehyde, acetone, or methanol fixation).

"Centromeres" and "kinetochores" refer to the site where chromosomes associate with the spindle during cell division. Direct visualization of ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function critically in cell motility and signaling. Since cilia are incapable of synthesizing their own protein, nearly 200 unique ciliary proteins need to be trafficked between the cytosol and primary cilia. However, it is still a technical challenge to map three-dimensional (3D) locations of transport pathways for these proteins in live primary cilia due to the limitations of currently existing techniques. To conquer the challenge, recently we have developed and employed a high-speed virtual 3D super-resolution microscopy, termed single-point edge-excitation sub-diffraction (SPEED) microscopy, to determine the 3D spatial location of transport pathways for both cytosolic and membrane proteins in primary cilia of live cells. In this article, we will demonstrate the detailed setup of SPEED microscopy, the preparation of cells expressing fluorescence-protein-labeled ciliary proteins, the real-time single-molecule tracking of individual proteins in live cilium and the achievement of 3D spatial probability density maps of transport routes for ciliary proteins.

Recently we mapped the three-dimensional (3D) spatial locations of transport routes for various proteins translocating inside primary cilia in live cells. Here this paper details the experimental setup, the process of biological samples and the data analyses for the 3D super-resolution fluorescence imaging approach newly applied in live primary cilia.

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function ...

Reconstruction of Single-Cell Innate Fluorescence Signatures by Confocal Microscopy

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing the innate cellular fluorescence signature from each individual live cell in a population distributed in a three-dimensional (3D) space. The innate fluorescence signature of a cell is a collection of fluorescence signals emitted by various biomolecules within the cell. Previous studies established that innate fluorescence signatures reflect various cellular properties and differences in physiological status and are a rich source of information for cell characterization and identification. Innate fluorescence signatures have been traditionally analyzed at the population level, necessitating a clonal culture, but not at the single-cell level. CRIF is particularly suitable for studies that require 3D resolution and/or selective extraction of fluorescence signals from individual cells. Because the fluorescence signature is an innate property of a cell, CRIF is also suitable for tag-free prediction of the type and/or physiological status of intact and single cells. This method may be a powerful tool for streamlined cell analysis, where the phenotype of each single cell in a heterogenous population can be directly assessed by its autofluorescence signature under a microscope without cell tagging.

Here, a protocol is presented for optically extracting and cataloging innate cellular fluorescence signatures (i.e., cellular autofluorescence) from every individual live cell distributed in a three-dimensional space. This method is suitable for studying the innate fluorescence signature of diverse biological systems at a single-cell resolution, including cells from bacteria, fungi, yeasts, plants, and animals.

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing ...

Customizable Fluorescence Microscope to Visualize Cytoskeleton Networks

Design and Building of a Customizable, Single-Objective, Light-Sheet Fluorescence Microscope for the Visualization of Cytoskeleton Networks

Reconstituted cytoskeleton composites have emerged as a valuable model system for studying non-equilibrium soft matter. The faithful capture of the dynamics of these 3D, dense networks calls for optical sectioning, which is often associated with fluorescence confocal microscopes. However, recent developments in light-sheet fluorescence microscopy (LSFM) have established it as a cost-effective and, at times, superior alternative. To make LSFM accessible to cytoskeleton researchers less familiar with optics, we present a step-by-step beginner's guide to building a versatile light-sheet fluorescence microscope from off-the-shelf components. To enable sample mounting with traditional slide samples, this LSFM follows the single-objective light-sheet (SOLS) design, which utilizes a single objective for both the excitation and emission collection. We describe the function of each component of the SOLS in sufficient detail to allow readers to modify the instrumentation and design it to fit their specific needs. Finally, we demonstrate the use of this custom SOLS instrument by visualizing asters in kinesin-driven microtubule networks.

Author Spotlight: Advancing Knowledge in Far-From-Equilibrium Materials Through Light-Sheet Microscopy

This protocol describes in detail how to build a single-objective, light-sheet fluorescence microscope and its usage for visualizing cytoskeleton networks.

Reconstituted cytoskeleton composites have emerged as a valuable model system for studying non-equilibrium soft matter. The faithful capture of the ...

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

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Chapters in this video

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

Super-resolution Imaging of the Bacterial Division Machinery

Quantitative Immunofluorescence Assay to Measure the Variation in Protein Levels at Centrosomes

Immunofluorescence Analysis of Endogenous and Exogenous Centromere-kinetochore Proteins

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

Reconstruction of Single-Cell Innate Fluorescence Signatures by Confocal Microscopy

Design and Building of a Customizable, Single-Objective, Light-Sheet Fluorescence Microscope for the Visualization of Cytoskeleton Networks